ISPM 27 27 ANNEX 18

ENG

DP 18: spp. INTERNATIONAL STANDARD FOR PHYTOSANITARY MEASURES PHYTOSANITARY FOR STANDARD INTERNATIONAL DIAGNOSTIC PROTOCOLS

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This diagnostic protocol was adopted by the Standards Committee on behalf of the Commission on Phytosanitary Measures in January 2017. The annex is a prescriptive part of ISPM 27.

ISPM 27 Diagnostic protocols for regulated pests

DP 18: Anguina spp.

Adopted 2017; published 2017

CONTENTS

1. Pest Information ...... 2 2. Taxonomic Information ...... 3 3. Detection ...... 4 3.1 Symptoms specific to Anguina ...... 4 3.1.1 (after Southey, 1972; Krall, 1991) ...... 4 3.1.2 Anguina (after Southey, 1973; Krall, 1991) ...... 4 3.1.3 Anguina funesta ...... 4 3.2 extraction ...... 5 3.2.1 Direct examination ...... 5 3.2.2 Extraction from soil and plant material ...... 5 3.2.3 Extraction from seed ...... 6 4. Identification ...... 6 4.1 Morphological identification ...... 6 4.1.1 Preparation of nematode specimens ...... 6 4.1.2 Morphological identification ...... 8 4.2 Molecular identification ...... 14 4.2.1 DNA extraction ...... 14 4.2.2 ITS1 rRNA PCR-RFLP for identification of Anguina spp. (Powers et al., 2001) ...... 15 4.2.3 TaqMan real-time PCR for identification of (Ma et al., 2011) ...... 17 4.2.4 Real-time PCR for identification of Anguina agrostis, A. funesta, A. pacificae and A. tritici (Li et al., 2015)...... 18 4.2.5 DNA sequence analysis of ITS1 and ITS2 rRNA ...... 20 4.2.6 Controls for molecular tests ...... 21 4.2.7 Interpretation of results from PCR ...... 22 5. Records ...... 22 6. Contacts Points for Further Information ...... 22 7. Acknowledgements ...... 23 8. References ...... 23 9. Figures ...... 27

International Plant Protection Convention DP 18-1 DP 18 Diagnostic protocols for regulated pests

1. Pest Information The family of contains both mycophagous species and species that parasitize bulbs, tubers and aerial parts of plants. The gall-forming nematodes of the subfamily Anguininae are obligate parasites of plants. More than 40 nominal species of gall-forming Anguinidae have been described. The seed gall nematodes, Anguina spp., inhabit the aerial parts of cereals and forage grasses. Species of Anguina invade ovules where they induce galls, lay eggs and reside as second-stage juveniles. This juvenile stage can remain as anhydrobiotes within dried seed galls for many years. A single wheat gall formed by Anguina tritici (Steinbuch, 1799) Filipjev, 1936 usually contains 11 000– 18 000 nematodes, although galls with as many as 90 000 have been recorded (Decraemer and Hunt, 2013). The nematodes can be retrieved from galls for up to 30 years after forming if kept dry. Three Anguina species, A. tritici (Steinbuch, 1799) Filipjev, 1936, A. agrostis (Steinbuch, 1799) Filipjev, 1936 and A. funesta Price, Fisher & Kerr, 1979 are considered of economic importance as agricultural and quarantine pests in various countries (Chizhov and Subbotin, 1990; Krall, 1991) This diagnostic protocol covers morphological and molecular identification of the and these species of major economic importance. Other species with importance in a limited geographical range include: A. agropyri Kirjanova, 1955 A. agropyronifloris Norton, 1965 A. amsinckiae (Steiner & Scott, 1935) Thorne, 1961 A. australis Steiner, 1940 A. balsamophila (Thorne, 1926) Filipjev, 1936 A. caricis Solovyeva & Krall, 1982 A. cecidoplastes (Goodey, 1934) Filipjev, 1936 A. graminis (Hardy, 1850) Filipjev, 1936 A. microlaenae (Fawcett, 1938) Steiner, 1940 A. pacificae Cid del Prado Vera & Maggenti, 1984 (Ferris, 2013). A. tritici (wheat gall nematode, bunted wheat) has been recorded in major wheat growing areas on all continents (Southey, 1972) and was historically widely distributed (EPPO, 2015). This species can cause severe crop losses to Secale cereale L. (rye) (35–65%) and Triticum aestivum L. (wheat) (20– 50%) (Leukel, 1929, 1957; Anwar et al., 2001). However, the use of modern seed cleaning methods that separate galls from healthy grains has almost eliminated this species from commercial wheat production in developed countries. For example, recent surveys for A. tritici in stored grain harvested from states of the United States of America with records of this nematode did not provide any evidence that it was still present in the country (CABI, 2001, 2014b). Recorded hosts are T. aestivum, Triticum dicoccum Schrank, Triticum durum Desf., Triticum monococcum L. (emmer), Triticum spelta L. (spelt), Triticum ventricosum Ces and S. cereale. Hordeum vulgare L. (barley) is a very poor host (Southey, 1972). There is little evidence that this nematode reproduces on Avena sativa L. (oat) and other grasses; although there are some reports of damage to oat at seedling stage by second-stage juveniles, no galls have been observed on this host. Clavibacter tritici, the bacterium that is the causal agent of yellow ear rot or “tundu” of T. aestivum, is associated with the presence of A. tritici. Freshly harvested infected wheat cockles containing the bacterium are toxic to cattle and sheep (Anwar et al., 2001). A. tritici has been shown to be vector of Rathayibacter toxicus (toxic yellow slime bacterium) under experimental conditions (Riley and McKay, 1990). A. agrostis (bentgrass nematode) has been reported from Asia, , Europe, New Zealand, North America and Republic of South Africa (CABI, 2014a). It is considered to be a species complex with pathotypes differing in host range (Krall, 1991; Brzeski, 1998). Subbotin et al. (2004) supported the concept of narrow specialization of seed gall nematodes, concluding that A. agrostis occurs in only

DP 18-2 International Plant Protection Convention Diagnostic protocols for regulated pests DP 18 one host, , and that other Agrostis species are hosts for two further undescribed species of Anguina. The type host of A. agrostis is Agrostis tenuis Sibth. In addition to various bentgrass species, A. agrostis sensu lato has been reported from other grass genera, including Apera, Arctagrostis, Calamagrostis, Dactylis, Eragrostis, Festuca, Hordeum, Koeleria, , Phalaris, Phleum, Poa, Puccinellia, Sporobolus and Trisetum (although certain records may relate to A. funesta (CABI & EPPO, 2004)). A. agrostis has been shown to vector R. toxicus under experimental conditions (Riley and McKay, 1990). Several older references (e.g. Goodey, 1960) to this species being the causal agent of disease in livestock relate to galls on Festuca spp. A. agrostis may actually refer to the species A. funesta (Southey, 1973). A. funesta has been described from Australia, and recently has been reported from Oregon in the United States of America (Meng et. al., 2012). The principal host of A. funesta is Lolium rigidum (annual ryegrass). A. funesta is recorded as a vector of R. toxicus, which causes the disease annual ryegrass toxicity when consumed by livestock. Annual ryegrass toxicity is responsible for severe losses in the livestock industry in Australia (Price et al., 1979). Rangeland infested by the nematode and bacterium is unusable for grazing (Figures 1 to 3).

2. Taxonomic Information Name: Anguina Scopoli, 1777 Synonyms: Angvina (= original spelling, amended to Anguina by later workers); Anguillulina Gervais & Van Beneden, 1859; Anguillulina (Anguina Scopoli) (Schneider, 1939); Paranguina Kirjanova, 1954; Paranguina Kirjanova, 1955 Taxonomic position: Nematoda, Chromadorea, Chromadoria, , Tylenchina, Tylenchomorpha, Sphaerularioidea, Anguinidae, Anguininae (after Decraemer and Hunt, 2013) Common names: Seed and leaf gall nematode, seed-gall nematode. Other common names in various languages are listed in CABI (2013). Name: Anguina tritici (Steinbuch, 1799) Chitwood, 1935 Synonyms: Vibrio tritici Steinbuch, 1799; Rhabditis tritici (Steinbuch) Dujardin, 1845; Anguillula tritici (Steinbuch) Grube, 1849; Anguillulina tritici (Steinbuch) Gervais & Van Beneden, 1859; tritici (Steinbuch) Bastian, 1865; Anguillula scandens Schneider, 1866; Tylenchus scandens (Schneider) Cobb, 1890; Anguillulina scandens (Schneider) Goodey, 1932 Common names: Wheat seed gall nematode, wheat cockle nematode Name: Anguina agrostis (Steinbuch, 1799) Filipjev, 1936 Synonyms: Vibrio agrostis Steinbuch, 1799; Anguillula agrostis (Steinbuch) Ehrenberg, 1838; Tylenchus agrostis (Steinbuch) Goodey, 1930; Anguillulina agrostis (Steinbuch) Goodey, 1932; Vibrio phalaridis Steinbuch, 1799; Anguilllula phalaridis (Steinbuch) Ehrenberg, 1838; Tylenchus phalaridis (Steinbuch) Örley, 1880; Anguillulina phalaridis (Steinbuch) Goodey, 1932; Anguina phalaridis (Steinbuch) Chizhov, 1980; Tylenchus agrostidis Bastian, 1865; Anguillula agrostidis (Bastian) Warming, 1877; Tylenchus phlei Horn, 1888; Anguina poophila Kirjanova, 1952; Anguina lolii Price, 1973

International Plant Protection Convention DP 18-3 DP 18 Diagnostic protocols for regulated pests

Common names: Bentgrass nematode, grass seed nematode Name: Anguina funesta Price, Fisher & Kerr, 1979 Synonyms: A. lolii Bird & Stynes, 1977 (in part) Common name: Seed gall nematode

3. Detection 3.1 Symptoms specific to Anguina species 3.1.1 Anguina tritici (after Southey, 1972; Krall, 1991) A. tritici incites seed galls (ear cockles) in cereals (Figure 4(A)). Invasive juveniles emerge from the seed galls in the soil and attack newly germinated seedlings. They establish infection on the tissues of young leaves near the growing point where they feed as an ectoparasite causing leaf distortion and crinkling (Figure 4(B)). Infected hosts become stunted and exhibit shorter and deformed stems and leaves (Figures 4(B) and 4(C)). Severely infected plants do not form ears or form only stunted ears on stunted stems. A diseased ear is much wider and shorter than a normal ear and has short deformed awns (Figure 4(C)). Slight elevations occur on the upper leaf surface with indentations on the lower side. Other leaf symptoms include wrinkling, twisting, curling of the margins towards the midrib, distortion, buckling, swelling and bulging. A tight spiral coil evolves, and dwarfing, loss of colour or development of a mottled yellowed appearance, and stem bending may also occur. In severe infection, the entire above-ground plant is distorted to some degree and therefore the disease is usually obvious (CABI, 2015). The second-stage juveniles stimulate the formation of galls in floral tissues in place of seed development. Galls vary from light and dark brown to almost black (Figure 5(A)). They are smaller than healthy grains (Figure 5(B)). The nematodes can survive in a quiescent stage in seed galls (Figure 6). 3.1.2 Anguina agrostis (after Southey, 1973; Krall, 1991) A. agrostis is considered to be an economically important nematode pest of bentgrass. In grasses, seed galls are difficult to detect as they are covered by lemmas and paleae. A small scarifier can be used to remove lemmas and paleae without damage to seeds or galls. This allows visual identification of galls (Alderman et al., 2003). Galled flowers have glumes of two or three times the normal length, lemmas five to eight times the normal length, projecting beyond the glumes as a sharp point, and paleae developing to about four times the normal length. Galls are at first greenish, and later become dark purple–brown. They reach 4–5 mm long (Figures 7 and 8). Lodicules, stamens and sometimes other flower parts are suppressed in parasitized flowers. Symptoms of the inflorescence also include elongated flower galls, which are modified ovaries that look greenish or purple and may be 4–5 mm long. Seed galls containing the nematodes are dark brown. They may look similar to normal seeds but are less heavy and hence can be separated mechanically from them. 3.1.3 Anguina funesta The life cycle of A. funesta is similar to that of A. agrostis. During dry summers, A. funesta survives within seed galls as anhydrobiotic second-stage juveniles. During winter, the nematodes are released from decaying galls and via water droplets in moist soil they invade new host seedlings, where they feed upon the young leaves. The nematodes congregate near the apical meristems until ovary initiation then stimulate ovary primordia to develop into galls. Occasionally, galls are produced in stamen primordia or, in very heavily infested plants, on glumes or rachis (McCay and Ophel, 1993). Information on the biology of this species can be found in Price et al. (1979). Symptoms of infestation are shown in Figures 1 to 3.

DP 18-4 International Plant Protection Convention Diagnostic protocols for regulated pests DP 18

3.2 Nematode extraction 3.2.1 Direct examination Symptomatic foliage and seed suspected to be infested with Anguininae can be processed by dissecting foliage tissue and galls immersed in tap water in a Petri dish. Specimens of motile and immotile stages may be observed under a stereomicroscope, usually within 30 min if the host plant is heavily infested. Seed cleaning can be achieved most effectively by modern equipment used for this purpose. Galls may be removed by a salt brine method in which the seeds are stirred into a 20% salt solution. Galls and debris float to the surface from where they are skimmed then steamed, boiled or chemically treated to kill the nematodes. The salt solution containing healthy seeds is drained and the seeds are rinsed several times in freshwater to remove the salt, then spread in thin layers on a clean surface to dry. 3.2.2 Extraction from soil and plant material Detailed descriptions of extraction equipment and procedures can be found in the European and Mediterranean Plant Protection Organization (EPPO) standard on nematode extraction (EPPO, 2013a). All stages of anguinid nematodes can be extracted from plant tissue, and infective juveniles can also be isolated from soil or growing medium, using the Baermann funnel technique, the modified Baermann tray method (Hooper and Evans, 1993), an adapted sugar flotation method (Coolen and D’Herde, 1972) or the mistifier technique (Hooper et al., 2005). These extraction methods should be conducted for 48 h at room temperature to detect low levels of infestation. Any plant material to be tested should be cut into pieces or sliced before extracting, for increased efficacy of extraction. The number of infective juveniles that may be recovered from soil depends on soil type, sampling depth, host plant and seasonal factors (Hooper, 1986). A large amount of fresh organic matter in the soil sample (e.g. plant residue after harvest) can influence nematode numbers because of its decomposition process, which might be toxic to nematodes or increase the number of saprophytic nematodes, or because the organic matter hampers extraction by clogging sieves or contaminating the supernatant obtained in density-based methods. The Baermann funnel technique (and modifications of it, such as the tray method, or Seinhorst mistifier, described by Hooper (1986)) is a reference technique for extraction of nematodes from soil and plant material. A piece of rubber tubing is attached to the stem of a glass funnel (with a recommended slope of approximately 30 degrees) and is closed by a spring or screw clip. The funnel is placed in a support and filled almost to the top with tap water. A plastic sieve or wire basket with a large enough aperture size to allow nematodes to actively pass through is placed just inside the rim of the funnel. Plant tissue cut into small pieces or soil is placed either directly onto the mesh or onto a single-ply tissue supported by the mesh, and the water level is adjusted so the substrate is only just submerged. Active nematodes pass through the mesh and sink to the bottom of the funnel stem. Alternatively, funnels made of plastic or stainless steel, or tubing made of silicone, can be used. However, regarding the latter, diffusion of oxygen into water is lower than for polyethylene (Stoller, 1957), which can lead to slow asphyxiation of the nematodes. Depending on the plant tissue, most (50–80%) of the motile nematodes present will be recovered within 24 h; however, samples can be left on the funnel for up to 72 h to increase the recovery rate. For longer extraction periods, regular tapping of the funnel and addition of freshwater increases nematode motility and compensates for evaporation and lack of oxygen, thereby improving the recovery rate. The efficacy of extraction can also be improved by adding 1–3% hydrogen peroxide (H2O2) for oxygen supply (Tarjan, 1967, 1972). Following the extraction period, a small quantity of water containing the nematodes is run off and observed under a stereomicroscope (Flegg and Hooper, 1970). Motile and immotile nematodes can be extracted from plant material by the sugar flotation method (Coolen and D’Herde, 1972). The plant material is washed, cut into pieces of about 0.5 cm, and 5 g portions are macerated in 50 ml tap water in a domestic blender at the lowest mixing speed for 2 min. The suspension of nematodes and tissue fragments is washed through a 750 µm sieve placed on top of

International Plant Protection Convention DP 18-5 DP 18 Diagnostic protocols for regulated pests a 45 µm sieve. The residue on the 45 µm sieve is collected and poured into two 50 ml centrifuge tubes. About 1 ml kaolin is added to each tube, the mixture is thoroughly stirred and then it is centrifuged at 1500 g for 5 min. The supernatant is decanted and sucrose solution (density 1.13 g/cm3) is added to the tubes. The mixture is thoroughly stirred and centrifuged at 1800 g for 4 min. The supernatant is washed through a 45 µm sieve, the residue is collected and the nematodes are studied under a stereomicroscope. Instead of sucrose, zinc sulphate (ZnSO4), magnesium sulphate (MgSO4) or colloidal silica can be used. 3.2.3 Extraction from seed Infective juveniles can be extracted from infested seed using a number of methods, including the Baermann funnel technique, which is summarized in section 3.2.2. A comparative study of the efficacy of various methods of extraction from seed, including water-agar blend, sieve blend, misting and blender-funnel-host stimulant, are presented by Griesbach et al. (1999). In the blender-funnel-host stimulant method, which is described in Griesbach et al. (1999) as the most effective method for the recovery of A. agrostis from Dactylis glomerata and Agrostis spp. seed, 50 g seed is placed in a blender with 300 ml tap water and blended for 15 s, shaken, and blended again for 15 s. The mixture is placed on a single tissue draped on an 850 μm pore size sieve supported over a large funnel containing tap water. Approximately 0.1 g orchardgrass leaves are added to the funnel as a stimulant, and the water column is aerated. Check valves at the funnel base enable the suspension to be drawn off after 24–48 h. The suspension is finally passed through a 25 μm pore size sieve before examination.

4. Identification The scope of this diagnostic protocol is to facilitate identification of Anguina to the genus level. Both morphological (section 4.1) and molecular (section 4.2) approaches are presented. 4.1 Morphological identification Information for morphological identification of valid genera within the Anguinidae is provided in section 4.1.2.1. Species of Anguinidae are probably one of the most variable groups among the Tylenchina regarding morphological characters (Brzeski, 1998). Identification to species can be unreliable if morphological characters are used in isolation; information regarding biology, host plant and symptoms of infection should also be taken into consideration. Often, only juveniles are found in seed galls, which can further complicate identification as important morphological features in adult specimens cannot be observed. Morphological information for the three species of economic importance is provided in section 4.1.2.2; however, this information should be used in combination with other sources to confirm diagnosis. Keys to species have been provided by Krall (1991) (ten species) and Brzeski (1998) (four species recorded from temperate Europe). 4.1.1 Preparation of nematode specimens As with other species of plant-parasitic nematodes, morphological observation should be carried out on as many adult specimens as possible. There are numerous published methods for fixing and processing nematode specimens for study, most recently summarized in Manzanilla-López and Marbán-Mendoza (2012). Nematodes processed with anhydrous glycerol are recommended for examination as important taxonomic features can be obscured if specimens are not cleared sufficiently. If possible, permanent slides should be prepared for future reference and deposited in nematode reference collections. Methods of preparing permanent slide mounts of nematodes have been described in detail elsewhere (Seinhorst, 1962; Hooper, 1986). The slow evaporation method as described by Hooper (1986), which preserves the structures and characteristics of the nematodes, is outlined in section 4.1.1.2. Temporary microscope slide preparations can be made quickly for instant examination but such slides may remain usable for only a few weeks.

DP 18-6 International Plant Protection Convention Diagnostic protocols for regulated pests DP 18

4.1.1.1 Temporary preparations A small drop of water is placed on a glass cavity slide, enough to sufficiently fill the well. The nematode specimens are transferred to the water and heated to 65 °C. It is critical that the heat should be applied just long enough to kill the nematodes, as prolonged heating will result in their distortion and deterioration. In practice, 10–15 s on a hotplate will be sufficient time for most species, but it is recommended that the slide be checked at intervals to monitor progress and removed from the heat only when movement of all the nematodes has ceased. A glass slide, free of dust, is placed on the side of the microscope stage. A small drop of single strength triethanolamine and formalin (TAF) fixative (10 ml formalin1 (35% formaldehyde in water) mixed with 1 ml triethanolamine and 89 ml distilled water) or another appropriate fixative is put in the centre of the slide and an appropriate amount of paraffin wax shavings is positioned around the drop (the wax will help support the coverslip and seal it to the slide). The nematodes are transferred from the cavity slide to the fixative so that they are positioned beneath the meniscus in the centre of the drop and not overlapping one another. The number of specimens able to fit on a slide will vary according to the size of the nematodes. An appropriately sized coverslip is carefully cleaned with lens tissue. It is gently lowered onto the wax shavings so that contact is made with the drop of fixative. The slide is placed on a hotplate and monitored until the wax has just melted; the air that may be lodged under the coverslip is removed by gently tapping the slide. The slide is then removed from the heat and examined. There should be a clear area of fixative containing the nematodes in the centre and a complete ring of wax to seal the slide. Should the seal be broken or the nematodes become embedded in the wax, the slide is heated again, the coverslip carefully removed, and the nematodes recovered and remounted on a new slide. If the wax has spread beyond the coverslip, it is cleared away with a fine blade. The coverslip is sealed with a ring of clear nail varnish. When the varnish has dried, the specimens are ready for study. 4.1.1.2 Permanent preparations A small drop of water is placed on a glass cavity slide, enough to sufficiently fill the well. The nematode specimens are transferred to the water and heated to 65 °C. It is critical that the heat should be applied just long enough to kill the nematodes, as prolonged heating will result in their distortion and deterioration. In practice, 10–15 s on a hotplate will be sufficient time for most species, but it is recommended that the slide be checked at intervals to monitor progress and removed from the heat only when movement of all the nematodes has ceased. The nematodes are transferred to an embryo dish or suitable watch glass half full of single strength TAF fixative (7 ml formalin1 (40% formaldehyde in water) mixed with 2 ml triethanolamine and 91 ml distilled water). The dish is covered and left to fix for a minimum of one week. The specimens are transferred to a watch glass containing a 3% glycerol solution with a trace amount of TAF fixative. The nematodes should be submerged. A coverslip is placed over the watch glass and left on it overnight. The coverslip is slightly moved so that a small gap is produced to allow evaporation, and the watch glass is left in an incubator (approximately 40 °C) until all the water has evaporated (this may take one week or longer). At the same time, a small beaker of glycerol is placed in the incubator to ensure it becomes anhydrous.

1 Formalin comprises 35–40% formaldehyde in water.

International Plant Protection Convention DP 18-7 DP 18 Diagnostic protocols for regulated pests

A small drop of the anhydrous glycerol is dispensed using a syringe or dropper onto the centre of a glass slide and the nematodes are transferred to this, arranged centrally. Three coverslip supports, such as glass beads, of similar diameter to that of the nematodes are placed at intervals in the margin of the glycerol drop so that they form an even support. Small amounts of paraffin wax shavings are placed at regular intervals around the circumference of the glycerol drop. A coverslip is heated on a hotplate for a few seconds. The coverslip is cleaned with lens tissue and gently lowered on to the wax, so that contact is just made between coverslip and glycerol. The slide is placed on the hotplate and as soon as the wax has melted and any air bubbles have been expelled by the settling coverslip, the slide is removed from the heat and the wax is allowed to reset. When the wax is completely hard, any excess wax is removed from around the coverslip with a scalpel. The coverslip is sealed with a ring of sealant such as Glyceel or clear nail varnish. The slide is labelled with an indelible marker, or affixed with a slide label. Information includes classification, date of slide preparation, host, locality, sample number (if applicable) and method of preservation. 4.1.2 Morphological identification 4.1.2.1 Morphological identification at the genus level Comparative morphology of genera assigned to the Anguininae is presented in Table 1. Definitions of terminology used in the following sections can be found in EPPO’s Diagnostic protocols for regulated pests: Pictorial glossary of morphological terms in nematology (EPPO, 2013b). Diagnosis of the Anguininae and the genus Anguina has been described by Siddiqi (2000), as follows and as described in Table 1, with key characters for identification shown in bold. Medium to large in size (1.0–2.7 mm), obese; mature female curved generally in one to one-and-a-half spirals. Metacorpus muscular. Basal bulb in adults enlarged, continuous or offset from isthmus by a constriction, base usually extending over anterior end of intestine. Ovary with one or two flexures anteriorly due to excessive growth; oocytes in multiple rows, arranged about a rachis. Crustaformeria a long tube formed by a large number of cells in multiple irregular rows. Spermatogonia in multiple rows. Bursa subterminal. Second-stage juveniles generally resistant, and the infective stage. Obligate plant parasites incite galls in seeds of cereals and grasses as well as stems, leaves and inflorescence of various monocotyledonous plants; type species causes wheat seed galls (ear cockles); only A. amsinckiae and A. balsamophila are known to parasitize dicotyledonous plants. There has been little recent morphological work on the genus and no reliable and up-to-date morphological keys to species are available. Therefore, identification at the genus level is described with summary information only for the three economically important pest species A. tritici, A. Agrostis and A. funesta.

DP 18-8 International Plant Protection Convention This diagnostic protocol was adopted by the Standards Committee on behalf of the Commission on Phytosanitary Measures in January 2017. The annex is a prescriptive part of ISPM 27.

Crustaform Incisures Female eria Post- Median Vulval Terminal in lateral Excretory Guber- Tail Genus Galls body Quadri- Rachis uterine pharyngeal Bursa flaps bulb fields pore naculum shape shape columella sac bulb

Litylenchus Zhao – Slender and + – – + +, non- Abuts 4 Posterior to + Extends to Conoid et al., 2011 semi-obese muscular nerve ring near tail tip Anguina Scopoli, + Obese, – + – + +, non- Base 4, unclear Posterior to + Small, Conoid 1977 spiral muscular or extends nerve ring subterminal muscular over intestine Diptenchus Khan, ? (roots) Slender + – – – +, muscular Abuts 5 At base of + To two-thirds Conical Chawla & Seshadri, pharynx tail length 1969 – (+ from Slender + – – + +, non- Abuts or 4 or 6 Posterior to + Adanal to Elongate Filipjev, 1936 India) muscular or overlaps a nerve ring subterminal conoid to muscular little filiform Indotylenchus ? Slender Not – – + +, muscular Offset 4 Anterior to + To two-fifths Elongate Sinha, Choudhury (mangroves) available median bulb tail length conoid & Baqri, 1985 Nothanguina + Obese, – + – + – Offset or Not Posterior to - To half tail Conoid Whitehead, 1959 spiral small dorsal available nerve ring length overlap Nothotylenchus – Slender + – – + +, non- Offset from 4 or 6 Posterior to + To half tail Elongate Thorne, 1941 muscular intestine nerve ring length conoid Orrina Brzeski, + Slender + – – + +, non- Overlaps a 4 Posterior to + To two-thirds Conoid 1981 muscular little nerve ring tail length PseudhalenchusT Not available Slender + – – + +, muscular Overlaps 4 Posterior to + To one-third Elongate arjan, 1958 nerve ring tail length conoid Pterotylenchus + Slender – – + + – Overlaps 4 Posterior to Not Not available Elongate Siddiqi & Lenné, dorsally nerve ring available conoid 1984 Safianema Siddiqi, – (fungal Slender + – – + +, muscular Overlaps 6 Posterior to + Adanal to Elongate 1980 feeder) laterally nerve ring subterminal conoid to filiform Subanguina + Slender or – – – + +, muscular Abuts or Not Posterior to + Subterminal Conoid Paramonov, 1967 semi-obese overlaps available nerve ring Zeatylenchus – Slender to + – – + +, non- Subventral Unable to In region of + Leptoderan, Conoid, Zhao et al., 2013 semi-obese muscular, glands discern in retracted toca 30% terminus fusiform overlap ♀, 3 at mid- stylet distance to with a intestine body in ♂ tail tip ventral spike Table 1. Comparative morphology and feeding habits of Anguininae

Source: After Zhao et al. (2011, 2013).

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4.1.2.2 Morphological identification of selected Anguina species Anguina tritici Description after Southey (1972) and Krall (1991). Refer to Table 2, and Figure 9 and Figure 10. Mature females. Body obese, spirally coiled ventrally. Lip region low and flattened, slightly offset, cephalic framework weak. Cuticle very finely annulated. Procorpus often swollen by gland secretions but constricted at junction of metacorpus. Isthmus sometimes posteriorly swollen, offset from pharyngeal glands by a deep constriction. Pharyngeal glands not overlapping, or with very slight overlap of intestine. Excretory pore near junction of pharynx and intestine or slightly more posterior. Vulval lips protruding, orifice of small glands visible on vulval lips anterior and posterior to vulva. Ovary with two or more flexures, often reaching to pharyngeal region with oogonia in multiple rows arranged around a rachis. Spermatheca pyriform, separated from oviduct by a sphincter. Postvulval uterine sac present. Tail conoid, tapering to an obtuse or rounded tip, not mucronate. Males. More slender than females. Habitus upon heat relaxation curved either ventrally or dorsally. Testis with one or two flexures. Spicules stout, arcuate, with two ventral ridges running from tip to widest part. Capitulum with distinct ventral folding at anterior. Gubernaculum simple, trough-like. Bursa leptoderan. Tail conoid, tip rounded or obtuse. Second-stage juveniles. Body slender, not spirally coiled. Tail conoid, pointed (Figure 10). For measurements, see Table 2. Anguina agrostis Description after Southey (1973) and Krall (1991). Refer to Tables 1 and 2, and Figures 11 and 12. Mature females. Based on specimens from the type host. Body obese, C-shaped to spirally coiled ventrally. Lip region low and flattened, offset by a fine constriction. Cuticle marked by fine annulations. Lateral fields not discernible on fully developed adult females, six incisures are visible on immature specimens. Neither procorpus nor isthmus exhibiting marked swellings, the former slightly contracted at its junction with the metacorpus. Isthmus occasionally folded in mature specimens by forward pressure of the gonad. Pharyngeal glands not overlapping, or with very slight overlap of intestine. Vulval lips prominent. Ovary usually with two flexures, often reaching to pharyngeal region with oogonia in multiple rows arranged around a rachis. Spermatheca pyriform, separated from oviduct by a constriction. Postvulval uterine sac present, 36–63% of vulva–anus distance. Tail conoid with an acute terminus. Males. Smaller and more slender than females. Habitus upon heat relaxation curved ventral to almost straight. Lateral field difficult to discern on mature specimens, reported to have six incisures. Testis usually reflexed once, spermatocytes arranged about a rachis. Spicules more slender in build than those of A. tritici, the capitulum showing little or no ventral folding at anterior. Two ventral ridges running from tip of each spicule to widest part, before converging and joining the capitulum. Gubernaculum simple, bursa leptoderan, ending just short of acute or finely mucronate tail tip. Second-stage juveniles. Body slender, not spirally coiled; tail conoid, pointed (Figure 12(A)). For measurements, see Tables 2 and 3. Anguina funesta Description after Price et al. (1979). Refer to Table 1 and Table 2, and Figure 13. Mature females. Length of postvulval uterine sac 62–112 µm; stylet length 7–10 µm. Young females are fully motile, but older females, in which gross expansion of the ovary has occurred, are strongly ventrally curved and capable of only weak movements of head and tail. Habitus following heat relaxation ventrally curved forming a complete circle, with head and tail overlapping. Lips slightly offset and rounded in front, cephalic framework lightly cuticularized. Stylet with conus and shaft of

DP 18-10 International Plant Protection Convention Diagnostic protocols for regulated pests DP 18 roughly equal length and with well-developed knobs. Pharynx 64–178 µm long with wide procorpus opening to a muscular metacorpus, ovate to spheroid, 17–25 µm long. Pharyngeal–intestinal junction obscured by the large dorsal gland, 45–72 µm in length, ovate to spatulate with a prominent nucleus. Pharyngeal glands slightly overlapping the intestine. Hemizonid between base of metacorpus and anterior end of dorsal pharyngeal gland, at 80–100 µm from anterior. Excretory pore located more posteriorly, 105–155 µm from anterior. Lateral field difficult to discern. Vulva with prominent lips. Anterior ovary with one or two flexures. Rarely, the gonad not reflexed but rather extended anteriorly to the base of the procorpus. Oocytes arranged in three or four rows about a rachis, except near the base of the ovary where this increases to five or six rows. Spermatheca ovate or elongated, long and 25–40 µm wide. Crustaformeria long and slender, made up of more than four columns of cells and up to 350 µm long, separated from spermatheca and uterus by short constrictions, 10–15 µm long. Crustaformeria often containing sperm cells and up to eight eggs. Uterus thick-walled, 70–100 µm long. Postvulval uterine sac approximately the same length as the uterus, 62–112 µm. Tail 48–112 µm long, vulva–anus distance 86–73 µm. Body width at anus approximately half that at vulva. Tail occasionally bluntly rounded, more usually conically pointed, sometimes with a mucronate tip. Males. Males shorter and thinner than females. Habitus upon heat relaxation straight or slightly curved. Lip and pharyngeal regions of the male similar to that of the female, although dorsal pharyngeal gland larger, almost rectangular, 50–78 µm long. Hemizonid and excretory pore 71–90 µm and 102–147 µm from anterior, respectively. Lateral field with five or six incisures, broken at intervals and occupying one-quarter to one-third of body width. Testis nearly always reflexed once. Testis with spermatocytes in multiple rows about a rachis. Spicules paired, non-fused and arcuate, each 16–28 µm long with characteristic bulges on the manubrium and where the manubrium and shaft join. Gubernaculum slim and trough-like. Bursa leptoderan, extending almost to tail tip, 44–114 µm in length. Tail 43–72 µm long with terminus conically pointed. Body width at cloaca 17–43 µm. For measurements, see Tables 2 and 3.

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Table 2. Morphometric data for Anguina tritici, Anguina agrostis and Anguina funesta Range, numerous populations Morphometric characters A. agrostis (restricted to A. tritici A. funesta Agrostis spp.) Mature females

L (mm) 3.0–5.2 1.3–2.7 1.65–2.44 Stylet (µm) 8–11 8–12 7–10 a (nematode body length/greatest 13–30 13.8–25.4 16.8–20.1 width (usually at mid-body)) b (nematode body length/pharynx length from lips to pharyngo- 9.8–25.0 8.0–28.7 9.3–34.0 intestinal valve) c (body length/tail length) 24–63 25.2–44.0 18.1–41.2 V (%) 70–95 87–92 86.9–94.0 Males L (mm) 1.9–2.5 1.05–1.68 0.78–1.52 Stylet (µm) 8–11 10–12 7–10 a (nematode body length/greatest 21–30 23–38 20.3–30.9 width (usually at mid-body)) b (nematode body length/pharynx length from lips to pharyngo- 6.3–13.0 6–9 6.3–9.5 intestinal valve) c (body length/tail length) 17–28 20.0–28.4 16.1–24.9 Spicules (µm) 35–40 25–40 16–28 Gubernaculum (µm) Approximately 10 10–14 9–14 Second-stage juveniles L (mm) 0.75–0.95 0.55–1.25 0.81–0.87 Stylet (µm) Approximately 10 Approximately 10 7–10 a (nematode body length/greatest 47–59 44–71 48–53 width (usually at mid-body)) b (nematode body length/pharynx length from lips to pharyngo- 4.0–6.3 3.2–6.1 4.2–4.6 intestinal valve) c (body length/tail length) 23–28 11.7–20 12.3–15.1

Source: After Southey (1972, 1973), Price et al. (1979), Chizhov (1980), Krall (1991), Brzeski (1998) and Meng et al. (2012). L, length; V, distance from the anterior end to the vulva divided by nematode body length (%).

Additional data for populations of A. agrostis and A. funesta (infective juveniles) are presented in Table 3.

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Table 3. Morphometric data of juveniles of an Anguina funesta population from annual ryegrass, an Anguina agrostis population from bentgrass and an Anguina agrostis population from orchardgrass from commercial seed production fields in Willamette Valley, Oregon, the United States of America Mean ± standard deviation (range) (µm) Characters A. funesta A. agrostis A. agrostis (n = 20) (annual (bentgrass) (orchardgrass) ryegrass)

836.2 ± 14.6 795.0 ± 33.8 739.8 ± 20.0 Body length (815.9–865.7) (726.4–875.6) (726.4–796.0) Genital primordium 396.7 ± 31.1 354.9 ± 25.0 350.0 ± 21.6 to posterior (351.5–480.2) (311.9–415.8) (297.0–381.2) 63.3 ± 3.3 57.7 ± 2.1 61.5 ± 4.2 Tail length (55.9–68.0) (53.5–60.8) (55.9–70.5) Anterior to 122.5 ± 2.5 122.2 ± 2.1 122.5 ± 2.5 excretory pore (119.1–128.8) (116.6–126.4) (119.1–128.8) 183.0 ± 7.1 186.9 ± 10.5 183.7 ± 8.8 Pharyngeal length (172.5–194.4) (158.0–199.3) (167.7–194.4) Genital primordium 20.4 ± 1.5 16.5 ± 1.5 18.1 ± 1.8 length (18.0–23.0) (13.5–19.0) (15.0–23.0) Genital primordium 8.8 ± 1.2 7.5 ± 0.9 8.1 ± 0.7 width (6.0–11.0) (6.0–10.0) (7.0–10.0) 16.6 ± 0.7 14.0 ± 0.5 14.6 ± 0.6 Body width (15.0–18.0) (13.0–15.0) (14.0–15.0) 17.0 ± 1.1 16.2 ± 1.6 15.9 ± 1.0 Metacorpus length (15.0–19.0) (12.0–19.0) (15.0–17.0) 8.5 ± 0.5 8.7 ± 0.8 7.9 ± 0.7 Metacorpus width (8.0–9.0) (7.5–10.0) (7.0–9.0) 8.0 ± 1.0 8.0 ± 1.0 8.0 ± 0.7 Stylet length (7.0–10.0) (7.0–10.0) (7.0–10.0) 10.1 ± 0.3 10.1 ± 0.5 10.5 ± 0.4 Anterior to base of stylet (10.0–11.0) (8.0–11.0) (10.0–11.0) 0.9 ± 0.4 1.2 ± 0.4 1.0 ± 0.3 Pharyngeal length (0.5–1.5) (0.5–1.5) (0.5–1.5) a (nematode body length/greatest width 50.4 ± 1.8 56.9 ± 3.7 50.6 ± 2.1 (usually at mid-body)) (48.0–52.9) (49.1–62.8) (45.4–54.7) b (nematode body length/pharynx length 4.6 ± 0.2 4.3 ± 0.2 4.0 ± 0.2 from lips to pharyngo-intestinal valve) (4.2–4.6) (4.0–4.7) (3.8–4.4) 13.2 ± 0.8 13.8 ± 0.6 12.1 ± 1.0 c (body length/tail length) (12.3–15.1) (12.8–15.0) (11.0–15.4) Anterior to excretory 14.6 ± 0.3 15.4 ± 0.6 16.6 ± 0.5 pore as % of length (14.4–16.0) (14.2–16.7) (15.3–17.1) Genital primordium 47.4 ± 3.6 44.7 ± 2.7 47.3 ± 3.0 to tail as % of length (43.2–56.8) (36.2–49.7) (39.8–51.8)

Source: Reproduced from Meng et al. (2012), courtesy Plant Management Network, Plant Health Progress.

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4.2 Molecular identification This section provides information on molecular tests that allow the identification of isolated nematodes of the major Anguina species. The tests are generally performed following a morphological examination in order to confirm the results obtained. Molecular diagnosis of Anguina spp. is based on polymerase chain reaction (PCR)-restriction fragment length polymorphism (RFLP) (Powers et al., 2001), real-time PCR (Ma et al., 2011; Li et al., 2015) or sequencing of the internal transcribed spacer (ITS) region of ribosomal (r)RNA (Subbotin et al., 2004). The choice of test depends on whether identification requires confirmation of both the presence and the absence of particular species, and on the availability of species standards for controls. The method described by Ma et al. (2011) is limited to positive identification of A. agrostis, while the other methods are able to simultaneously distinguish multiple species within the same test. PCR combined with analysis of RFLP is the most common way in which to simultaneously distinguish a range of Anguina species from each other (Powers et al., 2001). Powers et al. (2001) first sequenced the ITS1 region for Anguina spp. Subbotin et al. (2004) subsequently sequenced 58 populations of Anguina, Ditylenchus, Heteroanguina and Mesoanguina for phylogenetic analysis. There are 71 sequence accessions of rRNA fragments obtained from Anguina spp. collected from different localities and host plants presently available in the United States National Center for Biotechnology Information (NCBI) public database. ITS DNA PCR fragments may also be used for DNA sequence analysis, as described in section 4.2.5. In this diagnostic protocol, methods (including reference to brand names) are described as published, as these defined the original level of sensitivity, specificity and/or reproducibility achieved. The use of names of reagents, chemicals or equipment in these diagnostic protocols implies no approval of them to the exclusion of others that may also be suitable. Laboratory procedures presented in the protocols may be adjusted to the standards of individual laboratories, provided that they are adequately validated. 4.2.1 DNA extraction A single juvenile (or adult, if available) is processed for PCR by placing it in a 15 μl drop of double distilled sterile water on a glass slide and manually disrupting it by cutting it into several pieces with a knife under a stereomicroscope (Powers and Harris, 1993). The nematode pieces in 8 µl double distilled sterile water are transferred to a microcentrifuge tube containing, for example and depending on size and number of nematodes, 10 µl nematode extraction buffer (10 mM Tris, pH 8.2; 2.5 mM MgCl2; 50 mM KCl; 0.45% Tween 20; 0.05% gelatin; 60 µg/ml proteinase-K) (Thomas et al., 1997) and frozen at −70 °C for 15 min or until needed. The extract is thawed and incubated at 60 °C for 60 min then the proteinase-K is denatured by heating at 95 °C for 15 min. In the protocol by Ma et al. (2011) the nematode is cut in 8 µl double distilled water and this suspension is transferred to a tube containing 1 µl PCR buffer with 1 µl proteinase-K (1 µg/ml), with freezing as described above and incubation at 65 °C for 60 min followed by 95 °C for 10 min. There are no published protocols designed specifically for bulk DNA extraction from Anguina spp.; however, methods described for other nematodes can be adapted as required. For example, the commercially available QIAamp DNA Micro Kit (Qiagen2) was used for DNA extraction from reniform nematodes following Baermann extraction and sugar centrifugal flotation to isolate nematodes from soil (Sayler et al., 2012). Quantification of extracted DNA is measured with the NanoDrop ND-1000 spectrophotometer (Thermo Scientific2).

2 In this diagnostic protocol, methods (including reference to brand names) are described as published, as these defined the original level of sensitivity, specificity and/or reproducibility achieved. The use of names of reagents, chemicals or equipment in these diagnostic protocols implies no approval of them to the exclusion of others that may also be suitable. Laboratory procedures presented in the protocols may be adjusted to the standards of individual laboratories, provided that they are adequately validated.

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4.2.2 ITS1 rRNA PCR-RFLP for identification of Anguina spp. (Powers et al., 2001) The ITS rRNA universal primers described in this test are: rDNA2 (forward): 5′-TTGATTACGTCCCTGCCCTTT-3′ (Vrain et al., 1992) rDNA1.58S (reverse): 5′-ACGAGCCGAGTGATCCACCG-3′ (Cherry et al., 1997) The PCR and the cycling parameters as described by Szalanski et al. (1997) are presented in Table 4. Alternatively, the amplification can be conducted according to Meng et al. (2012) (Table 5). After PCR, 5 µl of the product is analysed electrophoretically on a 1.5% agarose gel in Tris-acetate- ethylenediaminetetraacetic acid (EDTA) (TAE) buffer. The gel can be stained with ethidium bromide and photographed using a gel imaging system with an ultraviolet light filter. The PCR products are purified with the Geneclean II Kit (MP Biomedicals2) or a similar PCR purification kit. The restriction enzymes AluI, BsrI, EcoRI, HaeIII, HhaI, HinfI and TaqI are required for identifying Anguina spp. The reactions are conducted separately (one tube for each enzyme) and according to the individual enzyme manufacturer’s recommendations (Table 6). A positive restriction control should be included in the RFLP step to confirm the success of the enzymatic digestion. The lengths of the restriction fragments generated by these diagnostic enzymes and the restriction pattern of each species are given in Table 7.

Table 4. ITS1 rRNA conventional PCR master mix composition, cycling parameters and amplicons (after Szalanski et al., 1997) Reagent Final concentration PCR-grade water –†

PCR buffer (including MgCl2) 1× dNTPs 0.8 mM Primer rDNA2 (forward) 0.4 mM Primer rDNA1.58S (reverse) 0.4 mM DNA polymerase 2.5 U DNA (volume) 1 µl Cycling parameters‡ Initial denaturation 94 °C for 3 min Number of cycles 40 Denaturation 94 °C for 45 s Annealing 55 °C for 1 min Elongation 72 °C for 2 min Final elongation 72 °C for 10 min Expected amplicons From 547 to 553 bp for Anguina spp. Size (except for Astrebla genus: 575 bp) † For a final reaction volume of 50 µl. ‡ According to the DNA polymerase manufacturer’s instruction. bp, base pairs; ITS, internal transcribed spacer; PCR, polymerase chain reaction; rRNA, ribosomal RNA.

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Table 5. ITS1 rRNA conventional PCR master mix composition, cycling parameters and amplicons (after Meng et al., 2012) Reagent Final concentration PCR-grade water –† PCR buffer 1×

MgCl2 1 mM dNTPs 0.4 mM Primer rDNA2 (forward) 0.2 mM Primer rDNA1.58S (reverse) 0.2 mM DNA polymerase (GoTaq Flexi (Promega2)) 2.0 U DNA (volume) 15 µl Cycling parameters‡ Initial denaturation 94 °C for 3 min Number of cycles 40 Denaturation 94 °C for 1 min Annealing 55 °C for 1 min Elongation 72 °C for 1 min Final elongation 72 °C for 10 min Expected amplicons From 547 to 553 bp for Anguina spp. Size (except for Astrebla genus: 575 bp) † For a final reaction volume of 50 µl. ‡ According to the DNA polymerase manufacturer’s instruction. bp, base pairs; ITS, internal transcribed spacer; PCR, polymerase chain reaction; rRNA/DNA, ribosomal RNA/DNA.

Table 6. Master mix composition, template, reaction conditions and amplicons for RFLP Reagent Final concentration PCR-grade water –† Enzyme mix 1× Restriction enzyme 10 U PCR product (volume) 6 µl Reaction conditions‡ 37 °C or 65 °C for 8 h Expected amplicons Size See Table 7 † For a final reaction volume of 14 µl. ‡ 37 °C for all enzymes except TaqI, for which incubation should be performed at 65 °C. PCR, polymerase chain reaction; RFLP, restriction fragment length polymorphism.

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Table 7. Restriction fragment sizes for Anguina species and associated restriction fragment length polymorphism (RFLP) patterns (after Powers et al., 2001) Species AluI * BsrI * EcoRI * HaeIII * HhaI * HinfI * TaqI * A. agrostis 548 A 295, B 299, 249 A 548 A 548 A 448, B 355, A 238, 15 100 135, 58 A. agropyronifloris 547 A 547 C 547 B 317, 230 B 547 A 447, B 489, 58 B 100 A. funesta 548 A 295, B 299, 249 A 548 A 548 A 448, B 490, 58 B 238, 15 100 A. graminis 548 A 310, B 548 B 548 A 548 A 249, C 490, 58 B 238 199, 100 A. microlaenae 550 A 550 C 301, 249 A 550 A 550 A 449, D 357, A 52, 49 135, 58 A. pacificae 549 A 239, D 549 B 319, 230 B 549 A 448, B 491, 58 B 225, 85 101 A. tritici 277, B 550 C 550 B 550 A 462, 88 B 550 E 492, 58 B 274 A. sp. / Dactylis 550 A 297, B 550 B 320, 230 B 550 A 252, C 357, A 253 198, 135, 58 100 A. sp. / Agrostis 553 A 301, B 553 B 333, 220 B 303, C 454, B 360, A 237, 15 250 99 135, 58 A. sp. / Polypogon 553 A 301, B 553 B 333, 220 B 553 A 454, B 360, A 237, 15 99 135, 58 A. sp. / Stipa 548 A 467, 81 A 299, 249 A 548 A 548 A 375, F 355, A 98, 46, 135, 58 29 A. sp. / Astrebla 359, C 575 C 575 B 575 A 575 A 401, A 517, 58 B 216 128, 46 A. sp. / Holcus 550 A 310, B 301, 249 A 550 A 550 A 450, B 303, C 240 100 135, 58, 54

* Code for the RFLP profile for each restriction enzyme. 4.2.3 TaqMan real-time PCR for identification of Anguina agrostis (Ma et al., 2011) This test developed by Ma et al. (2011) was designed as a species-specific real-time PCR to identify juveniles of A. agrostis. It was evaluated against the target species A. agrostis as well as non-target species A. tritici, A. wevelli and . While described for the “detection” of nematodes isolated from seed galls or plant material, the test was not specifically evaluated for its ability to quantify nematodes from quarantine samples. The ITS rRNA species-specific primers described in this test are: PF: 5′-GTTTGCCTACCGGTTGTTTACG-3′ PR: 5′-CCACATGCAGTCGGTGTGAA-3′ TaqMan probe Pb: 5′-FAM-TCATGTCTTGGCTATTGTAGACGTATCTGA-TAMRA-3′ The amplification is performed in a real-time PCR using the LightCycler (Roche2) according to the cycling parameters described in Table 8.

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Table 8. Real-time PCR master mix composition, cycling parameters and amplicons (after Ma et al., 2011) Reagent Final concentration PCR-grade water –† PCR buffer 1×

MgCl2 1.25 mM dNTPs 0.2 mM Primer PF (forward) 0.4 µM Primer PR (reverse) 0.4 µM Probe Pb 0.02 µM DNA polymerase 0.5 U DNA (volume) 1 µl Cycling parameters Initial denaturation 94 °C for 3 min Number of cycles 45 Denaturation 94 °C for 10 s Annealing 60 °C for 30 s Expected amplicons Size 88 bp † For a final reaction volume of 10 µl. bp, base pairs; PCR, polymerase chain reaction.

4.2.4 Real-time PCR for identification of Anguina agrostis, A. funesta, A. pacificae and A. tritici (Li et al., 2015) Li et al. (2015) designed a TaqMan real-time PCR to identify A. agrostis, A. funesta, A. pacificae and A. tritici. This test includes forward and reverse genus-specific primers combined with a fluorescent probe (modified with TET dye and BHQ-2 Black Hole Quencher2). This primers and probe set was designed to serve as an internal control for confirming the presence of Anguina spp. as well as the integrity of the PCR components and user performance. The test also includes primers and probe sets specifically designed for the detection of each of the target species mentioned above and is intended for identification of single juveniles. Species-specific probes were modified with 6-FAM and BHQ-1 and were simultaneously detectable on a different fluorescent channel in duplex PCRs (i.e. with the species-specific and genus-specific primers and probe sets). The sensitivity of the test was demonstrated through construction of standard curves from reactions using serially diluted nematode DNA: the test was able to detect as little as 1.25 copies of the ITS rDNA. The specificity of each primers and probe set was demonstrated in singleplex and duplex reactions (i.e. with the species- specific and genus-specific primers and probe sets) tested against all of the target species as well as several non-target nematodes including Anguina spp., Meloidogyne spp., spp. and Ditylenchus spp. The ITS rRNA genus- and species-specific TaqMan primers and probes described in this test are: A. agrostis (AAfpr primers-probe set) AAf (forward): 5′-CGGTTGTTTACGGCCGT-3′ AAr (reverse): 5′-ATGTAGTCGGTGTGAAAACAGCCAT-3′ AAp (probe): 5′-6-FAM/ATCATGTCTTGGCTATTGTAGACGTATCTG/BHQ-1-3′

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A. funesta (AFfpr primers-probe set) AFf (forward): 5′-GGTTGCTTACGGCCC-3′ AFr (reverse): 5′-GTGTAATCGATGTGATACAGCCCC-3′ AFp (probe): 5′-6-FAM/ATCATGTCTTGGCTATTATAGACGTATCTG/BHQ-1-3′ A. pacificae (APfpr primers-probe set) APf (forward): 5′-ACCGGTTGAATATTGGCTGT-3′ APr (reverse): 5′-ATGTAATCGATGTGAAACAGCCGT-3′ APp (probe): 5′-6-FAM/ATCATGTCTTGGAAAGTTTAGACGTATCTG/BHQ-1-3′ A. tritici (ATfpr primers-probe set) ATf (forward): 5′-GTTGCCTACGGCCGT-3′ ATr (reverse): 5′-ATGTAATCGATGTGGTACAGCCAT-3′ ATp (probe): 5′-6-FAM/ATCATGTCTTGGCTAGTGTAGACGTATCTG/BHQ-1-3′ Anguina spp. (ASfpr primers-probe set) ASf (forward): 5′-GTCTTATCGGTGGATCACTCGG-3′ ASr (reverse): 5′-TGCAGTTCACACCATATATCGCAG-3′ ASp (probe): 5′-TET/TCATAGATCGATGAAGAACGCAGCCA/BHQ-2-3′ The amplification reaction is performed in a real-time PCR using the SmartCycler II real-time PCR system (Cepheid2) according to the cycling parameters described in Table 9.

Table 9. Real-time PCR master mix composition, cycling parameters and amplicons (after Li et al., 2015) Reagent Final concentration PCR-grade water –†

PCR buffer (including MgCl2) 1×

MgCl2 6.0 mM dNTPs 0.24 mM Species-specific primer (forward) 240 nM Species-specific primer (reverse) 240 nM Species-specific probe 120 nM ASf internal control primer (forward) 160 nM ASr internal control primer (reverse) 160 nM ASp internal control probe 120 nM DNA polymerase (Platinum Taq (Invitrogen2)) 1.0 U DNA (volume) 1 µl Cycling parameters Initial denaturation 95 °C for 20 s Number of cycles 40 95 °C for 1 s Denaturation Optics OFF 60 °C for 40 s Annealing Optics ON Ramp 5 °C per s

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Expected amplicons Size 74–85 bp † For a final reaction volume of 25 µl. bp, base pairs; PCR, polymerase chain reaction.

4.2.5 DNA sequence analysis of ITS1 and ITS2 rRNA Molecular phylogenies of Anguina spp. provide a foundation for species identification based on alignment of rDNA sequences, including ITS1-partial 5.8S as described by Powers et al. (2001) or ITS1-5.8S–ITS2 as described by Subbotin et al. (2004). Sequences obtained from new isolates or unknown species are thus placed within the context of known species boundaries and phylogenetic relationships. For amplification of complete ITS1, 5.8S rDNA and ITS2, the primers used are: TW81 (forward): 5′-GTTTCCGTAGGTGAACCTGC-3′ (Joyce et al., 1994) AB28 (reverse): 5′-ATATGCTTAAGTTCAGCGGGT-3′ (Howlett et al., 1992) Alternatively, the following primer pair can be used: rDNA2 (also known as 18S) (forward): 5′-TTGATTACGTCCCTGCCCTTT-3′ (Vrain et al., 1992) rDNA1 (also known as 26S) (reverse): 5′-TTTCACTCGCCGTTACTAAGG-3′ (Vrain et al., 1992) The PCR is run with the composition and cycling parameters shown in Table 10. The PCR products are analysed by agarose gel electrophoresis. DNA fragments are extracted from the gel using commercially available reagents (e.g. Qiaquick Gel Extraction Kit (Qiagen2)) and sequenced with the same primers as for PCR. Alternatively, PCR products are cloned into a plasmid vector (e.g. TOPO TA cloning vector (Thermo Fisher Scientific2) or StrataClone vector (Agilent Technologies2)) and transformed into competent Escherichia coli. Plasmid clones are isolated from transformed bacteria using blue-white colony selection and sequenced using universal vector primers (Zheng et al., 2000). The size of the complete ITS1-5.8S–ITS2 region is approximately 675 base pairs for Anguina spp. DNA sequence alignment methods are numerous and rapidly evolving. DNA alignments with sequences obtained from GenBank are constructed with ClustalW, Clustal Omega or MAFFT from the European Bioinformatics Institute (available from http://www.ebi.ac.uk/Tools/msa) or by alignment plug-in modules within the commercial software packages Geneious (Biomatters2) and Chromas (Technelysium2). Pairwise genetic distances are calculated for all sequence combinations and expressed as percentage similarity or absolute number of nucleotide differences per aligned pair. Interspecific variation that exceeds the intraspecific variation generally indicates separation of species. A high degree of sequence similarity to named species should confirm results from PCR-RFLP and yield a definitive species diagnosis. A match to a previously identified unnamed population may yield the best possible conclusion for the circumstances.

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Table 10. DNA sequence analysis of ITS1 and ITS2 rRNA: PCR master mix composition, cycling parameters and amplicons Reagent Final concentration PCR-grade water –† Taq incubation buffer (Taq PCR Core Kit (Qiagen2)) 1× 5× Q-solution (Taq PCR Core Kit (Qiagen2)) 1× dNTPs 0.2 mM TW81 (forward) 1.5 µM AB28 (reverse) 1.5 µM DNA polymerase (Taq PCR Core Kit (Qiagen2)) 0.8 U DNA (volume) 10 µl Cycling parameters Initial denaturation 94 °C for 4 min Number of cycles 35 Denaturation 94 °C for 1 min Annealing 55 °C for 1 min 30 s Elongation 72 °C for 2 min Final elongation 72 °C for 10 min Expected amplicons Size 675 bp † For a final reaction volume of 100 µl. bp, base pairs; ITS, internal transcribed spacer; PCR, polymerase chain reaction; rRNA, ribosomal RNA.

4.2.6 Controls for molecular tests For the test result to be considered reliable, appropriate controls – which will depend on the type of test used and the level of certainty required – should be considered for each series of nucleic acid isolation and amplification of the target pest or target nucleic acid. For PCR a positive nucleic acid control, a negative amplification control (no template control) and a negative extraction control are the minimum controls that should be used. Positive nucleic acid control. This control is used to monitor the efficiency of the test method (apart from the extraction), and specifically the amplification. Pre-prepared (stored) nucleic acid of the target nematode may be used. Negative amplification control (no template control). This control is necessary for conventional PCR to rule out false positives due to contamination during preparation of the reaction mixture. PCR-grade water that was used to prepare the reaction mixture is added at the amplification stage. Negative extraction control. This control is used to monitor contamination during nucleic acid extraction. The control comprises nucleic acid extraction and subsequent amplification of extraction buffer only. It is recommended that multiple controls be included when large numbers of positive samples are expected. Positive digestion control (for RFLP only). This control is used to monitor the efficiency of the enzymatic digestion. Amplicon obtained from pre-prepared (stored) nucleic acid of target nematode may be used, as long as the nematode has been accurately identified as belonging to one of the species described in the RFLP patterns.

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4.2.7 Interpretation of results from PCR 4.2.7.1 Conventional PCR and PCR-RFLP The pathogen-specific PCR will be considered valid only if these criteria are met: - the positive control produces the correct size amplicon for the target nematode species - the negative extraction control and the negative amplification control produce no amplicons of the correct size for the target nematode species - the restriction enzyme patterns reveal only the bands expected for the species, with no additional bands and no missing bands. 4.2.7.2 Real-time PCR The real-time PCR will be considered valid only if these criteria are met: - the positive control produces an amplification curve with the species-specific primers and probe - the negative extraction control and the negative amplification control produce no amplification curve or no exponential curve - in the case of the Li et al. (2015) test, the genus-specific primers and probe produce an amplification curve in the presence of test sample DNA, indicating the presence of intact nematode DNA and integrity of the PCR components. A sample will be considered positive if it produces an exponential amplification curve. If a cycle cut- off value is needed, its value has to be verified in each laboratory when implementing the test for the first time.

5. Records Records and evidence should be retained as described in section 2.5 of ISPM 27 (Diagnostic protocols for regulated pests). In cases where other contracting parties may be affected by the results of the diagnosis, the following records and evidence and additional material should be kept for at least one year in a manner that ensures traceability: preserved or slide-mounted specimens, photographs of distinctive taxonomic structure, DNA extracts and photographs of gels. For morphological evidence, critical features as outlined in the morphological section should be drawn or photographed while fresh material is available, and relevant measurements should be included. Good photomicrographs (or scanning videos) of key morphological features are likely to be important for record keeping.

6. Contacts Points for Further Information Further information on this protocol can be obtained from: Nematology Laboratory, United States Department of Agriculture (USDA), Agricultural Research Service (ARS), 10300 Baltimore Ave., Bldg 010A BARC West, Rm 113, Beltsville, MD 20705, United States of America (Andrea Skantar; e-mail: [email protected]; tel.: +1 301 504 5917). Agri-Food and Biosciences Institute, Newforge Lane, Belfast BT9 5PX, United Kingdom (Colin Fleming; e-mail: [email protected]). Nematology Unit, Fera Science Limited, National Agri-Food Innovation Campus, Sand Hutton, York, YO41 1LZ, United Kingdom (Thomas Prior; e-mail: [email protected]; tel.: +44 1904 462206). A request for a revision to a diagnostic protocol may be submitted by national plant protection organizations (NPPOs), regional plant protection organizations (RPPOs) or Commission on

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Phytosanitary Measures (CPM) subsidiary bodies through the IPPC Secretariat ([email protected]), which will in turn forward it to the Technical Panel on Diagnostic Protocols (TPDP).

7. Acknowledgements The first draft of this protocol was written by Andrea Skantar (Nematology Laboratory, USDA-ARS, United States of America (see preceding section)), Colin Fleming (Agri-Food and Biosciences Institute, United Kingdom, Northern Ireland (see preceding section)) and Thomas Prior (Nematology Unit, Fera Science Limited, United Kingdom (see preceding section)).

8. References The present annex may refer to International Standards for Phytosanitary Measures (ISPMs). ISPMs are available on the International Phytosanitary Portal (IPP) at https://www.ippc.int/core- activities/standards-setting/ispms. Alderman, S.C., Bilsland, D.M., Griesbach, J.A., Milbrath, G.M., Schaad, N.W. & Postnikova, E. 2003. Use of a seed scarifier for detection and enumeration of galls of Anguina and Rathayibacter species in orchard grass seed. Plant Disease, 87: 320–323. Anwar, S.A., McKenry, M.V., Riaz, A. & Khan, M.S.A. 2001. Evaluation of wheat cultivars for Anguina tritici resistance, development and influence of nematode on wheat growth. International Journal of Nematology, 11: 150–156. Brzeski, M.W. 1998. Nematodes of Tylenchina in Poland and temperate Europe. Warsaw, Muzeum I Instytut Zoologii Polska Akademia Nauk. 395 pp. CABI. 2001. Anguina tritici. Crop Protection Compendium, global module, 3rd edn. Wallingford, UK, CABI. CABI. 2013. Datasheet for Anguina. Crop Protection Compendium. Wallingford, UK, CABI. Available at http://www.cabi.org/cpc (last accessed 6 March 2015). CABI. 2014a. Datasheet for Anguina agrostis. Crop Protection Compendium. Wallingford, UK, CABI. Available at http://www.cabi.org/cpc (last accessed 6 March 2015). CABI. 2014b. Datasheet for Anguina tritici. Crop Protection Compendium. Wallingford, UK, CABI. Available at http://www.cabi.org/cpc (last accessed 6 March 2015). CABI. 2015. Anguina species. Crop Protection Compendium. Wallingford, UK, CABI. Available at http://www.cabi.org (last accessed 25 April 2015). CABI/EPPO (European and Mediterranean Plant Protection Organization). 2004. Anguina agrostis. Distribution Maps of Plant Diseases, No. 923. Wallingford, UK, CABI. Cherry, T., Szalanski, A.L., Todd, T.C. & Powers, T.O. 1997. The internal transcribed spacer region of (Nemata: ). Journal of Nematology, 29: 23–29. Chizhov, V.N. 1980. The taxonomic status of certain species of the genus Anguina Scopoli, 1777. Byull Vesoyuznogo Instituta Gelmi-mintolii, No. 26: 83–95. Chizhov, V.N. & Subbotin, S.A. 1990. [Phytoparasitic nematodes of the subfamily Anguininae (Nematoda: ). Morphology, trophic specialization, taxonomic.] Zoologichesky Zhurnal, 69(4):15–26 (in Russian). Coolen, W.A. & D’Herde, C.J. 1972. A method for the quantitative extraction of nematodes from plant tissue. Ghent, Belgium, Ministry of Agriculture, State Agricultural Research Centre. 77 pp. Decraemer, W. & Hunt, D. 2013. Structure and classification. In R.N. Perry & M. Moens, eds. Plant nematology, 2nd edn, pp. 3–39. Wallingford, UK, CABI. 542 pp. EPPO (European and Mediterranean Plant Protection Organization). 2013a. Nematode extraction. Diagnostics PM 7/119 (1). EPPO Bulletin, 43: 471–495. EPPO (European and Mediterranean Plant Protection Organization). 2013b. Diagnostic protocols for regulated pests: Pictorial glossary of morphological terms in nematology. EPPO Technical

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Document No. 1056 (rev. 4). Paris, EPPO. 21 pp. Available at https://www.eppo.int/ QUARANTINE/diag_activities/EPPO_TD_1056_Glossary.pdf EPPO (European and Mediterranean Plant Protection Organization). 2015. EPPO Global Database. Paris, EPPO. Available at https://gd.eppo.int (last accessed 8 March 2015). Flegg, J.J.M. & Hooper, D.J. 1970. Extraction of free-living stages from soil. In J.F. Southey, ed. Laboratory methods for work with plant and soil nematodes, Technical Bulletin 2, pp. 5–22. London, Ministry of Agriculture, Fisheries and Food. 148 pp. Goodey, J.B. 1960. Gall-forming nematodes of grasses in Britain. Journal of the Sports Turf Research Institute, 10: 5–60. Griesbach, J.A., Chitambar, J.J., Hamerlynck, M.J. & Duarte, E.O. 1999. A comparative analysis of extraction methods for the recovery of Anguina sp. from grass seed samples. Journal of Nematology, 31(4S): 635–640. Hooper, D.J. 1986. Extraction of nematodes from plant tissue. In J.F. Southey, ed. Laboratory methods for work with plant and soil nematodes, Reference Book 402, 6th edn, pp. 51–58. London, Ministry of Agriculture, Fisheries and Food. 202 pp. Hooper, D.J. & Evans, K. 1993. Extraction, identification and control of plant parasitic nematodes. In K. Evans, D.M. Trudgill & J.M. Webster, eds. Plant parasitic nematodes in temperate agriculture, pp. 1–60. Wallingford, UK, CABI. 656 pp. Hooper, D.J., Hallmann, J. & Subbotin, S.A. 2005. Methods for extraction, processing and detection of plant and soil nematodes. In M. Luc, R.A. Sikora & J. Bridge, eds. Plant parasitic nematodes in subtropical and tropical agriculture, 2nd edn, pp. 53–86. Wallingford, UK, CABI. 871 pp. Howlett, B.J., Brownlee, A.G., Guest, D.I., Adcock, G.J. & McFadden, G.I. 1992. The 5S ribosomal RNA gene is linked to large and small subunit ribosomal RNA genes in the oomycetes, Phytophthora vignae, P. cinnamomi, P. megasperma f. sp. glycinea and Saprolegnia ferax. Current Genetics, 22: 455–461. Joyce, S.A., Reid, A., Driver, F. & Curran, J. 1994. Application of polymerase chain reaction (PCR) methods to identification of entomopathogenic nematodes. In A.M. Burnell, R.-U. Ehlers & J.P. Masson, eds. COST 812 Biotechnology: Genetics of entomopathogenic nematode- bacterium complexes. Proceedings of Symposium and Workshop. St Patrick’s College, Maynooth, Co. Kildare, Ireland, pp. 178–187. Luxembourg, European Commission, DG XII. 277 pp. Krall, E.L. 1991. Wheat and grass nematodes: Anguina, Subanguina and related genera. In W.R. Nickle, ed. Manual of agricultural nematology, pp. 721–760. New York, NY, Marcel Dekker. 1035 pp. Leukel, R.W. 1929. The nematode disease of wheat and rye. United States Department of Agriculture Farmers’ Bulletin, No. 1607. Leukel, R.W. 1957. The nematode disease of wheat and rye. United States Department of Agriculture Farmers’ Bulletin, No. 1607 (revised). Li, W., Zonghe, Y., Nakhla, M.K. & Skantar, A.M. 2015. Real-time PCR for detection and identification of Anguina funesta, A. agrostis, A. tritici, and A. pacificae. Plant Disease, http://dx.doi.org/10.1094/PDIS-09-14-0959-RE. Ma, Y., Xie, H., Wang, J. & Liu, C. 2011. Detection of second-stage juveniles of Anguina agrostis using TaqMan Real-time PCR. Russian Journal of Nematology, 19: 151–158. Manzanilla-López, R.H. & Marbán-Mendoza, N., eds. 2012. Practical plant nematology. Mexico City, Biblioteca Básica de Agricultura, Grupo Mundi-Prensa. 883 pp. McCay, A.C. & Ophel, K.M. 1993. Toxigenic Clavibacter/Anguina associations infecting grass seedheads. Annual Review of Phytopathology, 31: 151–167.

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Meng, S., Alderman, S., Fraley, C., Ludy, R., Sun, F. & Osterbauer, N. 2012. Identification of Anguina funesta from annual ryegrass seed lots in Oregon. Plant Health Progress, doi:10.1094/PHP-2012-1024-01-RS. Powers, T.O. & Harris, T.S. 1993. A polymerase chain reaction method for identification of five major Meloidogyne species. Journal of Nematology, 25: 1–6. Powers, T.O., Szalanski, A.L., Mullin, P.G., Harris, T.S., Bertozzi, T. & Griesbach, J.A. 2001. Identification of seed gall nematodes of agronomic and regulatory concern with PCR-RFLP of ITS1. Journal of Nematology, 33: 191–194. Price, P.C., Fisher, J.M. & Kerr, A. 1979. On Anguina funesta n. sp. and its association with Corynebacterium sp. in infecting Lolium rigidum. Nematologica, 25: 76–85. Pscheidt, J.W. & Ocamb, C.M., senior eds. 2015. Pacific Northwest plant disease management handbook (online). Corvallis, OR, Oregon State University. Available at http://pnwhandbooks.org/plantdisease (last accessed 9 November 2015). Riley, I.T. & McKay, A.C. 1990. Specificity of the adhesion of some plant pathogenic micro- organisms to the cuticle of nematodes in the genus Anguina (Nematoda: Anguinidae). Nematologica, 36: 90–103. Sayler, R.J., Walker, C., Agudelo, P. & Kirkpatrick, T. 2012. Conventional PCR detection and real-time PCR quantification of reniform nematodes. Plant Disease, 96: 1757–1762. Seinhorst, J.W. 1962. On the killing, fixing and transferring to glycerin of nematodes. Nematologica, 8: 29–32. Siddiqi, M.R. 2000. Tylenchida parasites of plants and insects, 2nd edn. Wallingford, UK, CABI. 864 pp. Southey, J.F. 1972. Anguina tritici. CIH description of plant parasitic nematodes, Set 1, No. 13. St Albans, UK, Commonwealth Institute of Helminthology. Southey, J.F. 1973. Anguina agrostis. CIH description of plant parasitic nematodes, Set 2, No. 20. St Albans, UK, Commonwealth Institute of Helminthology. Stoller, B.B. 1957. An improved test for nematodes in the soil. Plant Disease Reporter, 41: 531–532. Subbotin, S.A., Krall, E.L., Riley, I.T., Chizhov, V.N., Staelens, A., DeLoose, M. & Moens, M. 2004. Evolution of the gall-forming plant parasitic nematodes (Tylenchida: Anguinidae) and their relationships with hosts as inferred from Internal Transcribed Spacer sequences of nuclear ribosomal DNA. Molecular Phylogenetics and Evolution, 30: 226–235. Szalanski, A.L., Sui, D.D., Harris, T.S. & Powers, T.O. 1997. Identification of cyst nematodes of agronomic and regulatory concern by PCR-RFLP of ITS1. Journal of Nematology, 29: 255– 267. Tarjan, A.C. 1967. Influence of temperature and hydrogen peroxide on the extraction of burrowing nematodes from citrus roots. Plant Disease Reporter, 51: 1024–1028. Tarjan, A.C. 1972. Observations on extracting citrus nematode, Tylenchulus semipenetrans, from citrus roots. Plant Disease Reporter, 56: 186–188. Thomas, W.K., Vida, J.T., Frisse, L.M., Mundo, M. & Baldwin, J.G. 1997. DNA sequences from formalin-fixed nematodes: Integrating molecular and morphological approaches to . Journal of Nematology, 29: 250–254. University of Nebraska-Lincoln Nematology Lab. n.d. Plant and parasitic nematodes website. Anguina agrostis. Lincoln, NE, University of Nebraska-Lincoln. Available at http://nematode.unl.edu/aagrost.htm (last accessed 5 September 2016). Vrain, T.C., Wakarchuk, A.C., Levesque, A.C. & Hamilton, R.I. 1992. Intraspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fundamental and Applied Nematology, 15: 563–573. Zhao, Z.Q., Davies, K., Alexander, B. & Riley, I.T. 2011. Litylenchus coprosma gen. n., sp. n. (Tylenchida: Anguinata), from leaves of Coprosma repens (Rubiaceae) in New Zealand. Nematology, 13: 29–44.

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Zhao, Z.Q., Davies, K.A, Alexander, B.J.R. & Riley, I.T. 2013. Zeatylenchus pittosporum gen. n., sp. n. (Tylenchida: Anguinata), from leaves of Pittosporum tenuifolium (Pittosporaceae) in New Zealand. Nematology, 15: 197–212. Zheng, J., Subbotin, S.A., Waeyenberge, L. & Moens, M. 2000. Molecular characterisation of Chinese glycines and H. avenae populations based on RFLPs and sequences of rDNA-ITS regions. Russian Journal of Nematology, 8: 109–113.

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9. Figures

Figure 1. Healthy Lolium rigidum seed (left), Anguina funesta gall (centre) and nematode gall colonized by Rathayibacter toxicus (right). Photo courtesy I. Riley, South Australian Research and Development Institute (SARDI), Adelaide, Australia.

Figure 2. Healthy Lolium rigidum seed (left), Anguina funesta-infested nematode gall (centre) and Anguina funesta-infested bacterial gall (right). Photo courtesy J. Allen, Western Australia Department of Agriculture and Food, Perth, Australia.

Figure 3. Gumming disease of Lolium due to Rathayibacter toxicus. Photo courtesy J. Allen, Western Australia Department of Agriculture and Food, Perth, Australia.

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A B

C Figure 4. (A) Healthy Triticum aestivum ears (left) and ears infested with Anguina tritici (right). (B) and (C) Symptoms of infestation of T. aestivum seeds with A. tritici. Photos (A) © Howard Ferris, University of California, Davis, CA, United States of America, 1999; (B) courtesy Fera, United Kingdom; and (C) courtesy J. Swarup, India.

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Figure 5. (A) Healthy Triticum aestivum seeds (left) and seeds infested with Anguina tritici (right). Photo © Ulrich Zunke, University of Hamburg, Germany.

Figure 5. (B) Comparison of colour and shape of healthy Triticum aestivum seeds (left) and seeds infested with Anguina tritici (right). Photo courtesy T. Kościuch, Poland.

Figure 6. Invasive juveniles of Anguina tritici survive in a quiescent state within a seed gall, emerging to infest germinated seedlings. Photo © Howard Ferris, University of California, Davis, CA, United States of America, 1999.

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Figure 7. Agrostis plants infested with Anguina agrostis. Source: Pscheidt and Ocamb (2015, part of Ohio State University Extension Plant Pathology Slide).

Figure 8. Agrostis plants infested with Anguina agrostis. Photo © Malcolm Storey, 2011–2015.

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Figure 9. Anguina tritici from wheat grain: (A) vulval region showing a surface view of the uterus and postvulval uterine sac; (B) female; (C) oesophageal region of male; (D) head end of female; (E) male; (F) tail end of male; and (G) spicules. DN, nucleus of dorsal oesophageal gland; SVN, nuclei of subventral glands; RN, nucleus of renette cell. Reproduced from Siddiqi (2000), courtesy CABI.

Figure 10. Anguina tritici from wheat: (A) eggs and (B–E) second-stage juveniles showing (C) pharyngeal region, (D) tail and (E) lip region. Reproduced from Southey (1972), courtesy CABI.

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Figure 11. Anguina agrostis: (A) female; (B) male; (C) female pharyngeal region; (D) female tail; (E–F) male tails; and (G) spicule and gubernaculum. Reproduced from Southey (1973), courtesy CABI.

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A

B

C

Figure 12. Anguina agrostis juvenile: (A) whole nematode; (B) tail; and (C) head. Source: University of Nebraska-Lincoln Nematology Lab (n.d.). Photo © Peter Mullin, 2001.

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Figure 13. Anguina funesta: (A) adult male; (B) adult female; (C) male anterior; (D) male tail; and (E) spicule and gubernaculum. Reproduced from Price et al. (1979), courtesy Nematologica.

Publication history This is not an official part of the standard 2013-05 SC added subject Anguina spp. (2013-003) to the work programme under topic Nematodes (2006-008). 2015-03 Expert consultation on draft DPs. 2015-06 TPDP revised draft. 2015-11 TPDP e-decision for approval to SC (2015_eTPDP_Nov_01). 2015-12 SC approved draft for first consultation (2016_eSC_May_01). 2016-02 First consultation. 2016-10 TPDP recommended draft to SC for adoption (2016_eTPDP_Oct_01). 2016-10 SC approved draft to be submitted to the 45 day DP notification period (2016_eSC_Nov_04). 2017-01 SC adopted DP on behalf of CPM (no formal objections received). Publication history last updated: 2017-03

DP 18-34 International Plant Protection Convention This page is intentionally left blank IPPC The International Plant Protection Convention (IPPC) is an international plant health agreement that aims to protect cultivated and wild plants by preventing the introduction and spread of pests. International travel and trade are greater than ever before. As people and commodities move around the world, organisms that present risks to plants travel with them.

Organization ++ There are over 180 contracting parties to the IPPC. ++ Each contracting party has a national plant protection organization (NPPO) and an Official IPPC contact point. ++ Nine regional plant protection organizations (RPPOs) work to facilitate the implementation of the IPPC in countries. ++ IPPC liaises with relevant international organizations to help build regional and national capacities. ++ The Secretariat is provided by the Food and Agriculture Organization of the United Nations (FAO).

International Plant Protection Convention (IPPC) Viale delle Terme di Caracalla, 00153 Rome, Italy Tel: +39 06 5705 4812 - Fax: +39 06 5705 4819 Email: [email protected] - Web: www.ippc.int